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Dental pulp of the third molar: a new source ofpluripotent-like stem cells

Maher Atari1,2, Carlos Gil-Recio1, Marc Fabregat1, Dani Garcıa-Fernandez1, Miguel Barajas3,Miguel A. Carrasco4, Han-Sung Jung5, F. Hernandez Alfaro2, Nuria Casals6, Felipe Prosper3,Eduard Ferres-Padro2 and Luis Giner1,2,*1Laboratory for Regenerative Medicine, College of Dentistry, Universitat Internacional de Catalunya, Barcelona 08009, Spain2Surgery and Oral Implantology Department, College of Dentistry, Universitat Internacional de Catalunya, Barcelona 8017, Spain3Area of Haematology, University of Navarra, Pamplona 31008, Spain4Area of Pathology Universitat Internacional de Catalunya, Barcelona 08195, Spain5Division in Anatomy and Developmental Biology, Department of Oral Biology, Oral Science Research Center, College of Dentistry,Yonsei University, Seoul 120-749, South Korea6Basic Sciences Department and CIBER Physiopathology of the Obesity and Nutrition (CIBEROBN), Faculty of Medicine and Health Sciences,Universitat Internacional de Catalunya, Barcelona 08195, Spain

*Author for correspondence (lginer@csc.uic.es, drmaher1972@csc.uic.es)

Accepted 13 February 2012Journal of Cell Science 125, 3343–3356� 2012. Published by The Company of Biologists Ltddoi: 10.1242/jcs.096537

SummaryDental pulp is particularly interesting in regenerative medicine because of the accessibility and differentiation potential of the tissue.Dental pulp has an early developmental origin with multi-lineage differentiation potential as a result of its development duringchildhood and adolescence. However, no study has previously identified the presence of stem cell populations with embryonic-like

phenotypes in human dental pulp from the third molar. In the present work, we describe a new population of dental pulp pluripotent-likestem cells (DPPSCs) that were isolated by culture in medium containing LIF, EGF and PDGF. These cells are SSEA4+, OCT3/4+,NANOG+, SOX2+, LIN28+, CD13+, CD105+, CD342, CD452, CD90+, CD29+, CD73+, STRO1+ and CD1462, and they show genetic

stability in vitro based on genomic analysis with a newly described CGH technique. Interestingly, DPPSCs were able to form bothembryoid-body-like structures (EBs) in vitro and teratoma-like structures that contained tissues derived from all three embryonic germlayers when injected in nude mice. We examined the capacity of DPPSCs to differentiate in vitro into tissues that have similar

characteristics to mesoderm, endoderm and ectoderm layers in both 2D and 3D cultures. We performed a comparative RT-PCR analysisof GATA4, GATA6, MIXL1, NANOG, OCT3/4, SOX1 and SOX2 to determine the degree of similarity between DPPSCs, EBs and humaninduced pluripotent stem cells (hIPSCs). Our analysis revealed that DPPSCs, hIPSC and EBs have the same gene expression profile.

Because DPPSCs can be derived from healthy human molars from patients of different sexes and ages, they represent an easilyaccessible source of stem cells, which opens a range of new possibilities for regenerative medicine.

Key words: Dental pulp, DPPSC, Pluripotency, Teratoma formation, Embryonic markers, CGH technique

IntroductionStem cells have the ability to self-renew and to generate mature,

differentiated cells (Fuchs and Segre, 2000). The main postnatal

function of stem cells is to repair and regenerate the tissues in

which they reside. As pluripotent stem cells have become a major

focus of scientific research, many techniques have been developed

to determine the actual pluripotency of embryonic stem (ES) cells

or induced pluripotent stem (iPS) cells. The pluripotency of human

stem cells can be tested in two different ways: teratoma formation

by cells injected subcutaneously and the aggregation and

generation of embryoid bodies (EBs) from cells cultured in vitro

(O’Connor et al., 2008; Papapetrou et al., 2009). These techniques

demonstrate a multilineage differentiation capability by which

stem cells can give rise to cells of all three germ layers (Itskovitz-

Eldor et al., 2000; Martin and Evans, 1975).

Dental pulp tissue is thought to be derived from migratory

neural crest cells during development (Peters and Balling, 1999;

Thesleff and Aberg, 1999), and it has been shown to harbour

various populations of multipotent stem/progenitor cells (Miura

et al., 2003; Nosrat et al., 2001). To date, multiple human dental

stem/progenitor cells have been isolated, characterised, and

classified as a group designated ‘dental pulp stem cells’

(DPSCs). These include stem cells from exfoliated deciduous

teeth (SHEDs), periodontal ligament stem cells (PDLSCs), dental

follicle progenitor cells (DFPCs) and stem cells from apical papilla

(SCAPs). These post-natal populations have mesenchymal stem

cell (MSC)-like qualities, namely the capacity for self-renewal,

the potential to differentiate into multiple lineages, including

osteoblasts and chondroblasts, and a potential for in vitro

differentiation into cell types from various embryonic layers,

including adipose, bone, endothelial and neural-like tissues

(Arthur et al., 2008; Cheng et al., 2008; Cordeiro et al., 2008;

Fujii et al., 2008; Gay et al., 2007; Harada et al., 1999; He et al.,

2009; Honda et al., 2008; Huo et al., 2010). Many researchers have

proposed that DPMSCs are promising candidates for the repair and

regeneration of a variety of mesenchymal tissues, such as bone,

cartilage and muscle (Dezawa et al., 2005; Noel et al., 2002).

These findings, together with those of other studies, also suggest

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that cells from the dental pulp may represent a unique population

based on their regenerative potential (About et al., 2000; Gronthos

et al., 2000; Iohara et al., 2004; Mina and Braut, 2004; Zhang et al.,

2005). However, no optimal culture medium that allows adult stem

cell amplification without self-differentiation has yet been reported

(d’Aquino et al., 2007; Stevens et al., 2008).

No previous study to date has described the presence and

isolation of human dental pulp stem cells with pluripotent-like

characteristics at a single-cell level, nor the use of culture media

containing LIF (leukaemia inhibitor factor), EGF (epidermal

growth factor) and PDGF (platelet derived growth factor) for

isolation of these cells. In the present study, we describe the

isolation and identification of a subpopulation of pluripotent-like

stem cells from the dental pulp of third molars (termed DPPSCs)

that show greater regenerative power than currently used

mesenchymal stem cells. These DPPSCs are SSEA4+, OCT4+,

NANOG+, SOX2+, LIN28+, CD13+, CD105+, CD342, CD452,

CD90+, CD29+, CD732, STRO1+ and CD1462. We investigated

the capacity of DPPSCs to differentiate in vitro into tissues that

have similar characteristics to embryonic mesoderm, endoderm

and ectoderm layers in 2D and 3D, as well as their ability to

generate EB-like structures and to develop teratoma-like

structures when injected into nude mice. We also performed a

comparative analysis of GATA4, GATA6, MIXL1, NANOG,

OCT3/4, SOX1 and SOX2 by RT-PCR to determine the degree

of similarity between DPPSCs, EBs and human iPS cells.

DPPSCs are derived from an easily accessible source, and they

can be used in future protocols for the regeneration of tissues

from the three embryonic layers.

ResultsCharacterisation and isolation of dental pulp pluripotent

stem cells

These studies were carried out with the goal of isolating and

purifying a population of pluripotent-like stem cells derived from

dental pulp (DPPSCs). We analysed the phenotypes of all

populations, each of which corresponds to a dental pulp donor,

cultivated at a density of 802100 cells per cm2 and expanded at

different passages when the cultures reached 60% confluence.

Colony formation was observed, especially when the cells were

cultured by the hanging drop method. When colonies were

seeded into adherent surfaces cells tended to migrate

(supplementary material Fig. S1). We performed FACS, qRT-

PCR, immunophenotype analysis and cytogenetic analysis.

(Fig. 1A) shows representative images of DPPSCs morphology

at P5, P10 and P15. DPPSCs are small-sized cells with large

nuclei and low cytoplasm content, without the typical flat and

elongated MSC appearance. The morphology of DPPSCs

cultured in a Cell Carrier 3D glass scaffold was also examined

using scanning electronic microscopy (supplementary material

Fig. S2). Immunofluorescence assays for SSEA4, OCT3/4 and

NANOG showed that cells were positive for all three markers and

that SSEA4 localised in the cytoplasm, whereas embryonic

transcription factors were located in the nucleus (Fig. 1B).

The development of therapeutic strategies depends on the

ability of stem cells to undergo large scale in vitro amplification,

which can be associated with genetic instability. 85% of

DPPSCs exhibited a normal karyotype with no presence of

any aneuploidy, polyploidy or any chromosome structural

abnormality in metaphases after more than 65 passages (Fig. 1C).

Using transmission electron microscopy (TEM), we evaluatedthe morphology and integrity of the cells (Fig. 1D). A notable

feature of DPPSCs is that they possess large nuclei relative tocytoplasm volume, which is also a characteristic of ES cells.

Short-CGH analysis demonstrates genetic stability of DPPSCs

as they showed the same genetic dose as a healthy controlsample. A gain in X and a loss of Y chromosome dose can beseen, where this is due to sex differences: DPPSCs were extracted

from a female patient, whereas the cells onto which thehybridisation was performed were from a male donor (Fig. 1E).

In order to further characterise the population, we analysed the

cells by flow cytometry and found that the population wasCD105+, CD146+, CD452, CD342, STRO1+, TRA1-602, OCT3/4+ and NANOG2 (Fig. 2A,B). Double staining for OCT3/4 and

NANOG was also carried out showing a 19.55% of doublepositive cells (Fig. 2C). Some differences in the expression levelwere found between different passages. Interestingly, percentage

of SSEA4+, OCT3/4+ and NANOG+ increased with passages inDPPSCs, whereas in DPMSCs, they remained negative(supplementary material Table S2). We also looked for cellsexpressing embryonic markers in pulp tissues from donors of

different ages on the day of extraction (supplementary materialFig. S3). The percentage of SSEA4+ cells increased with age,whereas the number of OCT3/4+ and CD13+ cells decreased with

age. In addition, we found that embryonic markers were stillexpressed in DPPSCs from 58-year-old patients.

RNA was isolated from DPPSCs at P15, and expression of

OCT3/4, SOX2 and TERT was analysed by RT-PCR (Fig. 2D).Western blot analysis of OCT3/4 expression was performed inDPPSCs at P5, P10, P15 and P20. NTERA cells and bone marrow

multipotent adult progenitor cells (BM-MAPCs) were used aspositive controls, and Schwann cells and DPMSCs were used asnegative controls (Fig. 2E). Expression of OCT3/4 in DPPSCs

was maintained until at least P20.

The pluripotency of DPPSCs was assessed in vivo by teratomaformation. The injection of DPPSCs (P15) into nude mice

resulted in the formation of teratoma-like structures thatcontained tissues derived from all three embryonic germ layers.DPPSCs from two different donors gave similar results. DPMSCs

from the same donors were used as negative controls, and did notgive rise to teratoma formation (Fig. 2F). We performed theteratoma assays with four groups with a total number of seven 8-week-old nude mice (Samtako Bio Korea, Seoul, Korea) were

anesthetized with diethyl ether. Group 1:2 mice injected withcells from the 14-year-old donor. Group 2:2 mice injectedwith cells from the 17-year-old donor. Group 3:2 mice

injected with cells from the 28-year-old donor. Group 4:1 miceinjected with Matrigel (BD). Four out of seven mice injecteddeveloped teratoma-like forms with sizes of 0.6 to 1 cm just in

the left side, where DPPSCs were injected (two mice from group1, and two from group 3). In contrast, when DPMSCs from thesame donor were injected, no teratoma formation was observed

(Fig. 2E). All the mice from group 2 died at 8 or 12 days afterinjection. This result was unexpected and no obvious explanationoccurred to us.

Staining with H&E showed the formation of multiple adultstructures with origins in different embryonic layers (Fig. 2G;supplementary material Fig. S4) such as chondroid tissue,

chondroid matrix, fibroblasts and collagen fibres, adipose tissueand endothelium (Fig. 2L), gut-like epithelium (Fig. 2M), andneural-like tissue such as nerve and keratin (Fig. 2N).

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Immunohistochemical staining was performed to evaluate the

expression of embryonic markers after 3 weeks (Fig. 2H–K).

Although DPPSCs expressed embryonic markers when they were

undifferentiated, expression of these genes was lost during

differentiation, and very few cells were positive for embryonic

markers at 3 weeks. Antihuman antibodies were used to confirm

that the tissues formed were of human origin.

The ability of DPPSCs to form EBs was studied using a micro-

patterned culture surface and centrifugal force (Fig. 3A). After 5

days of culture, the morphology of EBs was evaluated by light

microscopy (Fig. 3B). EBs exhibited the typical spherical and

well-limited appearance of EBs formed from ES cells. Alkaline

phosphatase (ALP) staining was performed to confirm the

stem-like properties of the EBs (Fig. 3C). Furthermore, the EBs

continued to express embryonic markers such as OCT3/4 and

NANOG at day 5, as observed by immunofluorescence

(Fig. 3D,E). The expression of embryonic markers and lineage

specific markers was studied by RT-PCR. The results showed

that DPPSCs and EBs from DPPSCs expressed embryonic

markers such as OCT3/4, NANOG and SOX2 as well as other

lineage markers as SOX1, BDNF, MIXL1, GATA4 and GATA6

with levels comparable to iPS cells. DPMSCs did not express

these markers (Fig. 3F). To confirm the results, a qRT-PCR was

performed to check the expression of the same genes, using iPS

cells as a positive control. Levels of OCT3/4 and NANOG were

higher in DPPSCs than in EBs whereas lineage markers as GATA

4, GATA6 and MIXL1 were higher in EBs (Fig. 3G).

The protein expression profile of DPMSCs (Fig. 4A) was

substantially different from that of DPPSCs. Specifically, levels

of embryonic markers (OCT3/4, NANOG and SSEA4) were very

low, whereas levels of CD73 and STRO1 were high in DPMSCs.

Comparative FACS analysis was carried out on different

passages of populations from both cell types isolated from 14-,

17-, 18-, 28- and 38-year-old donors (supplementary material

Fig. 1. Characterisation and cellular morphology of

pluripotent stem cells obtained from dental pulp (DPPSCs) by

in vitro expansion. (A) Morphology of DPPSCs at different

passages (P5, P10 and P15). (B) Analysis of DPPSC immuno-

phenotype by confocal microscopy shows the expression of

SSEA4 PE (2861.13%) together with OCT3/4 FITC (60.365.3%)

or NANOG FITC (21.3%63.6). Average of three independent

experiments. (C) Cytogenetic analysis of undifferentiated DPPSCs

(P15) show 46 XY without aneuploidy or polyploidy;

chromosome structural abnormalities were not detected. (D) Cells

examined by transmission electron microscopy show large nuclei

and small cytoplasmic volume. Scale bar: 10 mm. (E) Short-CGH

analysis showing genetic stability of DPPSCs from a female

donor (n53).

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Table S2). The two cell types were easily distinguishable by their

morphology. DPMSCs showed the typical flat and elongated

appearance of mesenchymal cells, whereas DPPSCs were smaller

and more spherical in shape (Fig. 4B). Cell diameters rangedfrom 8–12 mm for DPPSCs and 12–19 mm for DPMSCs

(Fig. 4C).

Multiscreen-MIC plate carbonate filters were used to evaluate

the migratory capacity of DPPSCs. Although they came from the

same donor and were from the same passage, more DPPSCs

migrated across the filters than DPMSCs (Fig. 4D). To comparethe adhesive ability of DPPSCs and DPMSCs, the expression of

integrin CD29 was evaluated by FACS analysis. The results

showed that 99.6% of DPPSCs expressed integrin CD29,

compared to only 82.3% of DPMSCs (Fig. 4E).

Although they were the main population, DPPSCs coexisted in

culture with other cell types, such as mesenchymal stem cells

from the dental pulp (DPMSCs), due to the lack of a stringent

selection when performing primary culture and successivecultures with DPPSC medium. The two populations were

indistinguishable from each other by FACS analysis, although

size was known to be different. The percentages of OCT3/4,

NANOG and CD73 in different gates depending on size andcomplexity were insufficient to distinguish the two populations as

well (Fig. 4F,G; supplementary material Table S3).

Fig. 2. FACS characterisation, gene

expression of P15 DPPSCs and

teratoma formation. (A) FACS analysis

of P15 DPPSCs for membrane markers:

CD105 (90.7762.28%), CD146

(13.1760.68%), CD45 (0.0260.39%)

and CD34 (0.0660.05); (B) FACS

analysis of P15 DPPSCs for nuclear

markers: STRO1 (4.4261.23), TRA1

(0.0060%), OCT3/4 (62.9865.22%)

and NANOG (22.9662.58%).

(C) Analysis by FACS of DPPSCs with

double staining (19.55%) for OCT3/4

FITC (27.08%) and NANOG PE

(25.9462.51). Expressed as average of

three independent experiments. (D) RT-

PCR of OCT3/4, SOX2, TERT and

GAPDH of DPPSC P15 in 2D culture.

(E) OCT3/4 detection by western blot of

DPPSCs at different passages compared

with DPMSCs. NTERA cells and BM-

MAPCs were used as positive controls,

and Schwann cells and DPMSCs were

used as negative controls.

(F) Transplantation of P15 DPPSCs and

DPMSCs isolated from the same donor

from different ages (14, 17, 28 years old)

into immunodeficient mice resulted in

apparent teratoma-like formation (left

flank) after 5 weeks; DPMSCs P15 were

used as a negative control (right flank).

H&E staining of teratoma-like structures

induced by DPPSCs show the presence

of tissues from mesoderm, endoderm and

ectoderm. (G) H&E staining of teratoma-

like structure after 3 weeks. Circle

highlights zone that is magnified in H–K.

(H) Immunohistochemical staining for

OCT3/4. (I) Immunohistochemical

staining for SSEA4.

(J) Immunohistochemical staining for

NANOG. (K) Immunohistochemical

staining for LIN28.

Immunohistochemical staining shows

embryonic markers SSEA4, LIN28,

OCT3/4 and NANOG (black arrows).

(L) H&E staining showing chondroid

tissue, chondroid matrix, fibroblasts and

collagen fibres. (M) H&E staining

showing gut-like epithelium. (N) H&E

staining showing nerve-like tissue

and keratin.

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To further characterise the two cell populations, we performed

magnetic separation using a human PE selection kit. Cells

positive for CD73 were stained with a PE-conjugated antibody,

and we extracted RNA from both the CD732 and CD73+

populations after separation. RT-PCR was performed to

determine embryonic gene expression. The CD732 population

expressed TERT, OCT3/4 and SOX2, but the CD73+ cells did not

(Fig. 4H). This confirmed our previous assumption that DPPSCs

coexisted with DPMSCs when cultured in vitro.

We also observed a correlation between the embryonic

development stages of the third molar and the percentage of

OCT3/4 and NANOG expression by FACS analysis in pulp

tissues from donors of different nolla stages (6–10) and different

ages on the same day of extraction (n514 samples)

(supplementary material Table S4). To determine the

relationship between specific DPPSCs markers and tooth nolla

stages, the expression profiles were subjected to regression

analysis using nolla stage as the independent variable to

determine the subpopulation of DPPSCs. All the samples

showed the presence of DPPSCs and low expression of the

markers OCT3/4 and NANOG.

To see whether the culture conditions were a key aspect for the

maintenance of the different phenotypes between DPPSCs and

DPMSCs, we cultured each type of cell with the medium of the

other one. After one week changing the medium every 2 or 3

days we observed some phenotypic changes (Fig. 5A). DPPSCs

acquired a longer and flattered shape whereas some of the

DPMSCs became smaller and with a morphology resembling

DPPSCs. Changes were easier to see in DPPSCs culture in

mesenchymal media than in the other way. We checked the

expression of the embryonic markers that differ between the two

cell types. We observed that DPPSCs cultured in mesenchymal

medium lost the expression of NANOG, whereas the OCT3/4

levels only decreased. In the case of DPMSCs cultured in DPPSC

Fig. 3. Generation of DPPSC P15 embryoid bodies. (A) AggreWell system (Stem Cell Technologies), which utilises a micropatterned culture surface and

centrifugal forced aggregation to direct the formation of EBs by DPPSCs for 5 days. Scale bar: 100 mm. (B) Morphology of DPPSC embryoid bodies examined by

light microscope. (C) Alkaline phosphatase staining of DPPSC embryoid bodies. Scale bar: 100 mm. (D) Immuno-phenotype analysis by fluorescence microscopy

shows the expression of OCT3/4 FITC. (E) Immunostaining of DPPSC EBs generated by AggreWell system with NANOG FITC. (F) RT-PCR analysis of OCT3/

4, NANOG, SOX2, SOX1, BDNF, MIXL1, GATA4, GATA6 and GAPDH, comparing expression of hIPSCs, EBs from P15 DPPSCs and P15 DPMSCs and DPPSCs.

(G) qRT-PCR analysis of GATA4, GATA6, MIXL1, NANOG, OCT3/4, SOX1 in DPPSCs P15, EBs from DPPSCs and hiPSC. Expression levels were normalized to

GAPDH. Data represented as means 6 s.d. of three independent experiments.

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medium, cells gained the expression of both OCT3/4 and

NANOG (Fig. 5B).

To confirm the pluripotent capacity of DPPSCs we performed

three in vitro differentiation assays in which DPPSCs were

induced to give rise tissues from all three germ layers.

Mesoderm differentiation

We analysed the ability of DPPSCs to differentiate into osteoblasts

in 2D by immunofluorescence and qRT-PCR. The morphology of

DPPSCs cultured with osteogenic media changed over three weeks

of differentiation, resulting in cells with a bone-like appearance

(Fig. 6A) that expressed the osteoblast marker OSTEOCALCIN

(Fig. 6B). The differentiated cells showed upregulated expression

of specific bone tissue genes, such as ALP, OSTEONECTIN,

OSTECALCIN (Fig. 6C), OSTEOPONTIN, COLLAGEN I,

COLLAGEN III and BMP2, whereas NANOG was downregulated

(supplementary material Table S5). Human bone cDNAs were used

to normalise the data, GAPDH was used as a housekeeping gene

and DPPSC undifferentiated dental pulp cells were used as a

negative control (data not shown). ALP, OSTEONECTIN and

Fig. 4. Comparison of morphology and protein profiles in DPPSCs and DPMSCs. (A) Immuno-phenotype by FACS analysis of P15 DPMSCs isolated from

dental pulp. Expression of STRO-1 (84.1261.67%), NANOG (0.2860.33%), SSEA4 (0.0960.12%), OCT3/4 (0.260.2%), CD73 (7262.69%) is shown as the

average of three independent experiments. (B) Comparison of morphology of DPPSCs and DPMSCs from the same donor and the same passage (P15). (C) Cell

diameters of DPPSCs and DPMSCs measured by Scepter Millipore. (D) Cell migration capacity of P15 DPPSCs and DPMSCs from the same donor and hNTERA

cells, which were used as a positive control, was determined using a Multiscreen-MIC Plate polycarbonate filter (8 mm pore size), incubated at 37 C, 5% CO2 for 6

hours. Significance was set at *P#0.05 (n55). (E) Analysis by FACS for CD29 expression in a 3D culture of DPPSCs at P15 (99.660.61%) and DPMSCs at P15

(82.367.88%). (F,G) Analysis of the different populations coexisting in DPPSC cell culture in terms of their cell size (FSC) and their complexity (SSC).

Percentage of OCT3/4+, NANOG+ or CD73+ cells in every gate analysed is shown with respect to the total. (H) RT-PCR for embryonic genes TERT, OCT3/4 and

SOX2 of the two separated cell populations of DPPSC (CD732) and DPMSC (CD73+) cultures.

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OSTEOCALCIN expression increased in the first, second and third

weeks of differentiation. We also observed a decrease in NANOG

expression. To confirm bone-like cell differentiation, we used

Alizarin Red to stain extracellular matrix deposits consisting of

hydroxyapatite, calcium and magnesium salts (Fig. 6D). Taken

together, these assays demonstrate that DPPSCs could efficiently

differentiate into bone-like tissue and express specific bone tissue

genes.

Differentiation was also performed using a Cell Carrier 3D

glass scaffold. When cultured in osteogenic differentiation

medium, DPPSCs derived into bone-like tissue that was able to

synthesise typical bone structures, such as collagen and cortical

structures that were detectable by SEM analysis (Fig. 6E;

supplementary material Fig. S5). The 3D differentiation was

also confirmed by qRT-PCR. During three weeks of

differentiation, ALP, COLLAGEN I and COLLAGEN III

expression steadily increased, whereas the embryonic markers

NANOG and OCT3/4 decreased each week (Fig. 6F). Functional

activity was determined by quantifying ALP activity (Fig. 6G)

and the calcium secretion (Fig. 6H) every week for three weeks.

Both ALP activity and the calcium secretion increased

significantly on days 7, 14 and 21 of osteogenic differentiation.

To demonstrate the capacity of DPPSC to differentiate into

other tissues of the mesoderm cap, we performed a vessel-derived

endothelial cell differentiation, in which DPPSCs were cultured

with basal media (2% FBS, 50 ng/ml VEGF, 10 ng/ml bFGF) for

Fig. 5. Changes in expression of embryonic markers upon changing the

culture conditions. (A) Changes in morphology in DPPSCs and DPMSCs

when they were cultured in the medium of the other cell type for 1 week.

(B) Changes in expression of embryonic markers in P10 DPPSCs and

DPMSCs when cultured in the medium of the other cell type for 1 week

(2 passages). Expression of OCT3/4 and NANOG was analysed by RT-PCR.

Fig. 6. Differentiation of P15 DPPSCs into mesoderm. (A) Cell morphology

during osteogenic 2D differentiation at weeks 1, 2 and 3 (W1, W2 and W3).

(B) Immunofluorescence shows the expression of osteocalcin FITC in the

cytoplasm and cell mask Alexa Fluor W696. (C) ALP, OSTEONECTIN and

OSTEOCALCIN detection by qRT-PCR at different time points during 2D

osteogenic differentiation of DPPSCs (n53, from 14-, 17- and 28-year-old

donors, *P,0.05). The mRNA levels were normalised to GAPDH (a

housekeeping gene). The relative expression was normalised to human cDNAs,

which is normalised to 1. (D) Alizarin Red staining of DPPSCs after 3 weeks of

2D osteogenic differentiation. (E) Scanning electron microscopy image of 3D

differentiation of DPPSCs at P15 using a Cell Carrier 3D glass scaffold for 21

days into mesoderm tissues. Bone-like tissue, collagen (white arrow) and

cortical (green arrow) structures were observed. (F) Analysis of the expression

of ALP, COLLAGEN I, COLLAGEN II, NANOG and OCT3/4 by qRT-PCR

during 3 weeks of 3D differentiation. The mRNA levels were normalised to

GAPDH. The relative expression was normalised to human cDNA from bone

(n53, from 14-, 17- and 28-year-old donors, *P,0.05, **P#0.001), which was

designated as 1. (G) ALP activity at different time points of osteogenic

differentiation in 3D for 21 days. Data are presented as means 6 s.d. (n53).

*P#0.05. (H) Calcium quantification at different time points of osteogenic 3D

differentiation for 21 days. Data are presented as means 6 s.d. (n53). *P#0.05.

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three week. We observed an increase in the expression of specific

endothelial genes, such as FLK1 and CD14 (by qRT-PCR) and

FLK1 (by immunofluorescence) (supplementary material Fig. S6).

Endoderm differentiation

Another differentiation assay was performed to induce DPPSCs

to generate cells from the endodermal lineage, specifically

hepatocyte-like cells. DPPSCs cultured in 2D with hepatogenic

media for the three weeks showed altered morphologies, with

some cells adopting a polygonal shape similar to hepatocytes

(Fig. 7A). Expression of the typical hepatic protein albumin

(ALB) was detected by immunofluorescence (Fig. 7B). Using

qRT-PCR, we demonstrated that differentiated cells expressed

the hepatic nuclear factors HNF3b, HNF6 and GATA4 (Fig. 7C).

The expression levels of a-fetoprotein (AFP), ALB, CEBPA and

NANOG were also determined (supplementary material Table

S6). We also observed a decrease in NANOG during the 3 weeks

of differentiation (data not shown). Human liver cDNAs

(Ambion) were used to normalise the data and GAPDH was

used as the housekeeping control (data not shown). The levels of

secreted ALB increased significantly over three weeks of

differentiation (Fig. 7D). These results indicate DPPSCs can

upregulate endoderm markers and differentiate into hepatic-like

cells, both genetically and functionally, upon hepatogenic

induction.

Cell Carrier 3D glass scaffolds were also used to perform the

endoderm differentiation. After 21 days of 3D culture with

hepatogenic media, some hepatic structures were seen by SEM

analysis, including large and small pores with a fenestra-like

appearance, the surfaces of sinusoidal endothelial cells and ultra-

structures of sinusoidal endothelial cells (Fig. 7E; supplementary

material Fig. S7). qRT-PCR was performed to evaluate the

expression of hepatic specific genes ALB, AFP and CYP3A4.

Expression of these markers increased each week, whereas the

embryonic markers NANOG and OCT3/4 expression decreased

(Fig. 7F). Functional activity was determined by quantifying

Fig. 7. Differentiation of P15 DPPSCs into

endoderm. (A) Cell morphology of DPPSCs at

different time points (weeks 1, 2 and 3) during 2D

hepatic differentiation. (B) Immunofluorescence at day

21 of 2D differentiation into hepatocyte-like cells shows

the expression of ALB FITC in the cytoplasm and cell

mask Alexa Fluor W696. (C) qRT-PCR detection of

GATA4, HNF3b and HNF6 in 2D differentiated

DPPSCs (n53, *P,0.05). The mRNA levels were

normalised to GAPDH. The relative expression was

normalised to human cDNAs from liver cells, assigned

as 1. (D) ALB synthesis analysis at different days of 2D

differentiation into hepatocyte-like cells. Data are

presented as mean 6 s.d. (n53 from 14-, 17- and 28-

year-old donors). *P#0.05. (E) Scanning electron

microscope image of 3D endoderm differentiation of

DPPSCs at P15 after 21 days on a Cell Carrier 3D glass

scaffold. Hepatic-like cells with large and small open

pores are observed that have a fenestrated appearance

(white arrow). The ultrastructure of sinusoidal

endothelial cells (green arrow) and the surface of

sinusoidal endothelial cells can also be observed.

(F) qRT-PCR detection of ALB, AFP, CYP3A4,

NANOG and OCT3/4 in DPPSCs differentiated into a

3D hepatic lineage (n53, *P,0.05, **P#0.001). The

mRNA levels were normalised to GAPDH. The relative

expression was normalised to human cDNAs from liver

cells, which were assigned as 1. (G) ALB synthesis

analysis at different days of 3D differentiation into

hepatocyte-like cells. Data are presented as means 6

s.d. (n53). *P#0.05. (H) Cytochrome P450-3A4

Metabolic Activity Assay. Data are presented as means

6 s.d. (n53). *P#0.05.

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ALB secretion (Fig. 7G) and P450 CYP3A4 activity (luciferin-

PFBE) (Fig. 7H). As in 2D differentiation cultures, the levels of

secreted ALB increased over time. The activity of CYP3A4 in

differentiated cells was higher than in undifferentiated control

cells. Both experiments were performed in triplicate.

Ectoderm differentiation

Finally, DPPSCs were able to follow a neuroectodermal

differentiation pathway in 2D cultures. The morphology of

differentiated cells was phenotypically clearly similar to neuronal

cells (Fig. 8A). Immunofluorescence staining showed that

differentiated DPPSCs expressed neuronal tissue-specific proteins

such as NEUN (Fig. 8B), GALC 595 and PSD95 (Fig. 8C). qRT-

PCR analysis showed that expression levels of TAU increased with

time, and that the levels of NURR1 and NESTIN increased during

weeks one and two and then plateaued at week three (Fig. 8D).

GFAP, MBP and SOX1 also increased during the first, second and

third weeks, whereas NANOG expression decreased during

differentiation (supplementary material Table S7). Human brain

cDNA (Ambion) was used to normalise the data and GAPDH was

used as the housekeeping control (data not shown).

DiscussionSeveral populations of stem cells have been isolated from

different parts of the human tooth and all of them have been

shown to have generic mesenchymal stem cell-like properties

(Laino et al., 2005). These cells express markers associated withthe endothelium and/or the smooth muscle, such as Stromal-

derived factor 1 (STRO1), Vascular cell adhesion molecule 1(VCAM1), Melanoma-associated antigen MUC18 (MUC18) andsmooth muscle actin (Tecles et al., 2005). Other reports havedescribed the presence of a lateral population of stem cells in the

dental pulp (Casagrande et al., 2006; Liu et al., 2006; Rizzino,2009; Sloan and Smith, 2007; Volponi et al., 2010; Zhang et al.,2006b). However, there has been no previously published

reference to the presence of a population of cells in dental pulpwith the protein profile SSEA4+, OCT3/4+, NANOG+, Nestin+,SOX2+, LIN28+, CD13+, CD105+, CD342, CD452, CD90low,

CD29+, CD73low, STRO1low and CD1462 (Oda et al., 2010), aspresented in this work. OCT3/4, NANOG and SOX2 areindispensable for indefinite stem cell division, without affectingdifferentiation potential or the capacity for self-renewal. The

functional importance of SOX2 and NANOG genes in altering theprogenitor status has also been clearly demonstrated (Hanna et al.,2010; Ratajczak et al., 2008; Takahashi and Yamanaka, 2006; Yu

et al., 2007; Zuba-Surma et al., 2009). NANOG has been reportedto be a key gene for maintaining pluripotency, as shown by thecapacity for multilineage differentiation and perpetual self-

renewal of cells expressing this gene. Although DPPSCs sharesome features with other populations of stem cells in the tooth,they differ in other aspects, such as gene expression and

differentiation potential (Hirata et al., 2010) due to theirembryonic-like properties. In this work, we observed theformation of EB-like structures with characteristics similar toembryonic stem cells and MAPCs (Braccini et al., 2005;

Verfaillie, 2005).

Both DPMSCs and DPPSCs are obtained from dental pulpusing the same isolation protocol. Distinction of these different

cell populations is dependent on the density at which the cells areseeded and the culture medium. The use of media that containgrowth factors EGF, PDGF and LIF, allows maintenance of the

pluripotent state of DPPSCs. There has been no other report ofthe use of a similar medium to culture stem cells from the dentalpulp, so it seems likely that this media formulation is key formaintaining DPPSCs with typical stem cell properties. In culture,

however, DPPSCs display heterogeneity that can be explained asa consequence of their spontaneous differentiation and the lack ofclonality upon their isolation. As previously shown, other stem

cell cultures undergo the same processes; for example, MAPCcultures are heterogeneous, and two cell populations withdifferent phenotypes (large and small) coexist in the same

culture (Verfaillie, 2005). Here, we have demonstrated that theDPPSC population coexists in culture with other cell types, suchas DPMSCs, and that the populations were different from one

another, as they differed in pluripotency gene expression aftermagnetic separation by CD73. It seems plausible that both celltypes have a common progenitor, but the relationship betweenthem is still unclear. The fact that DPPSCs express some

embryonic markers but the other population does not suggeststhat these populations may be at a different level of thedifferentiation hierarchy, and that DPPSCs may be the

progenitors that gave rise to the other populations. Furtherexperiments will be needed in order to test this hypothesis.

The third molars are the most common source of dental stem

cells, because wisdom tooth extraction is widely performed andthe teeth are usually considered to be medical waste (Otaki et al.,2007; Yang et al., 2007). Because the third molar is the last tooth

Fig. 8. Neuroectoderm differentiation of P15 DPPSCs. (A) Image by

optical microscopy of the morphology of cells differentiated into neural-like

cells at different time points (weeks 1, 2 and 3). (B) DPPSCs differentiated

into neural-like cells after 3 weeks showed expression of NEUN FITC and

cell mask Alexa Fluor W696. (C) DPPSCs differentiated to neural-like cells

after 3 weeks showed expression of GALC595 and PSD95. (D) Analysis of

the expression of NURR 1, TAU and NESTIN by qRT-PCR during the three

weeks of differentiation. mRNA levels were normalised to GAPDH. The

relative expression was normalised to human cDNA from brain cells (n53,

from 14,- 17- and 28-year-old donors, *P,0.05), which was designated as 1.

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to develop in humans, it is normally in an early stage ofdevelopment and is capable of yielding an optimum quantity of

dental pulp tissue for the isolation of DPSCs (Gandia et al., 2008;Laino et al., 2006; Leeb et al., 2010; Morsczeck et al., 2009;Smith et al., 2009; Atari et al., 2011; Zhang et al., 2006a).Although the percentage of DPPSCs decreases with age, a

population of these cells was always present, even in olderpatients. Multipotent cells that have a certain degree ofpluripotency, because they are derived from early embryonic

cells and maintained in the adult stage (Ebert et al., 2009), havebeen found in adult tissues, such as very small embryonic-like(VSEL) stem cells in the bone marrow. These studies, together

with the data shown here, indicate that the dental pulp could bean important source of cells with pluripotent characteristics. Wespeculate that DPPSCs could be derived from residualundifferentiated cells in the dental pulp. However, this

hypothesis requires further testing. The characteristics unique tothese cells are still under investigation, but the current evidenceopens to the way for future comparative studies of the

regenerative potency of DPPSCs and stem cells from othersources. In addition, the possibility of freezing these cellsfollowing molar extraction seems feasible, as is currently

performed with cells obtained from the umbilical cord.

In this paper, we have shown that DPPSCs have pluripotent-like properties that have not been found in cells of any other adultsource to date. The ability to form EB-like or teratoma-like

structures has been thought to be exclusive to ES or iPS cells(Benton et al., 2009; Maltman and Przyborski, 2010). With thisfinding, a new field of investigation can be opened. If one

population of cells found in adult individuals can achieve truepluripotency, there may be other populations with the same orsimilar properties that have yet to be discovered. The relationship

between DPPSCs and iPS cells should also be investigated. Theinduction of iPS cells seems to be easier from stem cells thanfrom differentiated cells (Illich et al., 2011). It could be that a

reprogramming process occurs in DPPSCs, but not indifferentiated dermal fibroblasts. Thus, only DPPSCs couldselectively expand when cultured in DPPSC media, but othercells from the dental pulp could not undergo the reprogramming

process needed to acquire pluripotency. Importantly, the numberof reprogramming factors needed for induced pluripotency couldbe reduced when using DPPSCs. Further studies are needed to

answer these questions.

For therapeutic purposes, the reliability and safety of putativeclinical applications for DPPSCs must be considered, especially

the issue of genetic stability. We have demonstrated that DPPSCsshow no chromosome abnormalities when cultured in vitro, suchthat we propose that DPPSCs are safe to use for clinical therapies.We propose that short-CGH should be used in stem cell research

to determine genetic stability when cells are cultured in vitro,because s-CGH allows the detection of genetic abnormalities thatcould remain hidden with the current protocols, such as

karyotype or FISH techniques. In addition, stem cells fortherapeutic applications must be able to differentiate intodifferent tissues. As we have shown, DPPSCs are capable of

giving rise to mesodermal, endodermal and ectodermal tissuesthat express markers typical of osteoblasts (Atari et al., 2012),hepatocytes and neurons, respectively, as shown by qRT-PCR

assays, in which all data were normalised to human cDNAs of therespective tissues. The use of 3D culture systems to carry outdifferentiation protocols represents an improved system that

simulates the physiological in vivo environment (Dhawan et al.,

2010; Undale et al., 2009). Based on images obtained by inverted

optical microscopy (2D differentiation) and the pictures taken by

SEM (3D differentiation), we observed tissue-specific structures,

such as collagen fibres and fenestra-like structures, in 3D that

were not seen in 2D.

These cells have properties that are not observed in other cells

obtained from adult tissues, which could open a new range of

possibilities for regenerative medicine. The results presented here

suggest that DPPSCs may be useful for treating various disorders,

such as those related to the loss of bone or some of the proteins

synthesised related with the malfunction of liver cells. Further

investigations are needed to fully characterise these cells.

Materials and MethodsPatient selection

Healthy human third molars extracted for orthodontic and prophylactic reasons

were selected from 20 different patients of different sexes and ages (14–60 years

old). The extraction procedure was kept simple to prevent tooth damage. Dental

pulp tissues used for these experiments were obtained with informed consent from

donors. All experiments were performed in accordance with the guidelines on

human stem cell research issued by the Committee on Bioethics of the

International University of Catalonia.

Primary cells obtained from human molar samples

Immediately after extraction, the third molars were washed using gauze soaked in

70% ethanol, followed by a wash with sterile distilled water. Holding the tooth

with upper incisor forceps, an incision was made between the enamel and the

cement using a cylindrical turbine bur. A fracture was made on the same line, andfragments of the tooth were placed in a Falcon flask containing sterile 16 PBS.

The samples were rapidly transported to the laboratory and placed in Petri dishes in

a laminar flow hood. Tissues were isolated from the dental pulp using a sterile

nerve-puller file 15 and forceps. Cellular separation was completed by digesting

the divided pulp tissue with collagenase type I (3 mg/ml) (Sigma) for 60 minutes at

37 C. Cells were then separated using an insulin syringe and centrifuged for 10

minutes at 275 g (RCF). The cell fraction was washed twice with sterile 16PBS

and centrifuged again for 10 minutes at 275 g at room temperature. Once collected,the cells were counted and seeded in DPPSC medium. In order to establish the

primary culture, the cells were grown in 96-, 24- and 6-well culture dishes and in

150 ml flasks coated with 100 ng/ml human fibronectin inside a 5% CO2

humidified chamber for 3 weeks. The medium was changed every 4 days. During

the splitting/passaging of DPPSCs, cell density was maintained at 80–100 cells/

cm2 by detaching cells with 0.25% trypsin (Cellgro) and replating every 36–48

hours, when cells were 60% confluent.

DPPSC culture medium

The cell expansion medium consisted of 60% DMEM low glucose (Sigma) and

40% MCDB-201 (Sigma) supplemented with 16 Insulin-Transferrin-Selenium

(ITS) (Sigma), 16linoleic acid bovine serum albumin (LA-BSA) (Sigma), 1029 M

dexamethasone (Sigma), 1024 M ascorbic acid 2-phosphate (Sigma), 100 units ofpenicillin, 1000 units of streptomycin (PAA), 2% foetal bovine serum (Sigma),

10 ng/ml hPDGF-BB (R&D Systems), 10 ng/ml EGF (R&D Systems), 1000 units/

ml hLIF (Chemicon), Chemically Defined Lipid Concentrate (Gibco), 0.8 mg/ml

BSA (Sigma) and 55 mM b-mercaptoethanol (b-ME, Sigma).

Base medium

The base medium consisted of 60% DMEM low glucose (Gibco), 40% MCDB-201

(Sigma) with 16 Insulin-Transferrin-Selenium, 16 linoleic acid BSA, 1029 M

dexamethasone (Sigma), 1024 M ascorbic acid 2-phosphate (Sigma), 100 units of

penicillin and 1000 units of streptomycin (Gibco).

Isolation and culture of human dental pulp mesenchymal stem cells

(DPMSCs)

Human adult DPMSCs were isolated from the dental pulp of third molars and were

suspended in Dulbecco’s Modified Eagle’s Medium (DMEM, Biochrom) containing

2 ng/ml basic fibroblast growth factor (bFGF) and 10% foetal bovine serum (FBS,

Hyclone). Cells were plated at a density of 300,000 cells/cm2. The medium was

changed after 72 hours and every 2 days thereafter. To propagate DPMSCs, the cellswere detached at 90% confluence by the addition of phosphate buffered saline (PBS,

Biochrom) containing 0.05% trypsin-ethylenediaminetetraacetic acid (EDTA,

Biochrom) and replated at a density of 4000 cells/cm2.

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Human NTERA-2 and IPS

NTERA-2 cells were obtained from the ATCC. Cells were maintained and culturedin Dulbecco’s Modified Eagle’s Medium supplemented with 10% foetal bovineserum and 1% penicillin-streptomycin at 37 C humidified atmosphere at 5% CO2.hIPS wire was kindly donated by our collaborators at the University of Navarra.

Flow cytometry

FACS analysis was carried out the same day of the extraction and again after twoand three weeks of culture initiation. The following fluorochrome-labelledmonoclonal antibodies were used: CD13 FITC (eBioscience), SSEA4 PE(eBioscience), OCT3/4 FITC (RD SYSTEMS), CD45 PE-Cy5 (BDPharmingen), CD105 FITC (BD Pharmingen), CD34 PE-Cy5 (BD Pharmingen),CD73 PE (BD Pharmingen), CD146 FITC (BD Pharmingen), CD90 FITC(eBioscience), CD29 PE (BD Pharmingen), STRO1 FITC (BD Pharmingen),LIN28, SSEA1 PE, SOX2 PE and NANOG FITC (Abcam). For the analysis ofcontrol samples, different IgG isotypes coupled to FITC, PE and PE-Cy5fluorochromes (BD Pharmingen) were used. The cell suspension (in PBS plus 2%FBS) was incubated for 45 minutes at 4 C in the dark. Later, cells were washedtwice with PBS containing 2% FBS and centrifuged for 6 minutes at 275 g (RCF).Depending on the number, cells were resuspended in 300 to 600 ml of PBS and 2%FBS. All flow cytometry measurements were made using a FACScan cytometer(FACSCalibur) and analysed using the winMDI 2.8 program.

Karyotyping

The cells were trypsinised and centrifuged at 250 g (RCF). The cell pellet wasresuspended in a volume less than or equal to 500 ml of DMEM-LG and was thensubjected to a hypotonic shock with 0.075 M KCl that had been pre-heated to 37 C(the saline solution was added drop by drop under continuous agitation). After30 minutes of incubation in the presence of KCl at 37 C, the cells were centrifugedat 300 g (RCF) for 10 minutes. The nucleus suspension was fixed twiceconsecutively in methanol:acetic acid (3:1). After the final centrifugation step, thefinal cell pellet was resuspended in fixation liquid and extensions were performedon glass slides that were pre-cooled to 4 C. The extensions were stained usingGIEMSA stain and the number of cells in metaphase was counted using a lightmicroscope at a 1006 magnification with oil immersion. A minimum of 50metaphases were counted per sample.

RNA isolation and qRT-PCR

Total cellular RNA samples were extracted using Trizol (Invitrogen) from thefollowing cell types: DPPSCs at passages 5, 10 and 15 (P5, P10 and P15), H-NTERA, DPMSCs and differentiated cells. RNA was extracted weekly from thedifferentiated cells. Two mg of RNA was treated with DNase I (Invitrogen) andreverse-transcribed using M-MLV reverse transcriptase (Invitrogen). We analysedthe efficacy of the cDNA (1, 0.1, 0.01, 0.001, 0.0001 dilutions) at differentconcentrations for all primers of pluripotent genes using NTERA cells as positivecontrols. Additionally, we tested the following samples as positive controls:hepatocyte markers from human liver cDNA samples, osteoblast markers fromhuman bone cDNA and neuroectoderm markers from human brain cDNA(Ambion). Quantitative RT-PCR was performed using the CFX96 thermocycler(Bio-Rad). Quantitative RT-PCR was performed using 50 ng of cDNA and SYBR

Green Supermix (Bio-Rad Laboratories, Inc.). cDNA samples were amplifiedusing specific primers with the following conditions for 40 cycles. The expressionlevels of genes of interest (supplementary material Table S1) were normalisedagainst the housekeeping gene GAPDH. The relative expression levels werenormalised to human cDNAs (positive controls), which were assigned as 1. Theresults were analysed using the 2DDCt method.

Western blotting

Total protein was extracted from the following samples: DPPSCs collected atdifferent passages (P5, P10, P15 and P20), DPMSCs, NTERA-2, HEK 293 andbone marrow derived-MAPCs. Cell lysates with equal protein concentrations(20 mg/ml) were separated by SDS-PAGE on 12% polyacrylamide gels andtransferred onto nitrocellulose membranes. The membranes were blocked with 1%(w/v) BSA in PBS containing 0.1% Tween 20, blotted with OCT3/4 and GAPDHprimary antibodies (1:5000) and blotted with secondary antibodies (1:15,000). Allantibodies were purchased from Abcam.

Immunofluorescence analysis

Samples were fixed with 4% paraformaldehyde (Sigma) for 4 minutes at roomtemperature followed by methanol (Sigma) for 2 minutes at 220 C. For nuclearligands, cells were permeabilised with 0.1 M Triton X-100 (Sigma) for 10 minutes.Slides were incubated sequentially for 30 minutes each with primary antibody andFITC, PE or PE-Cy5-coupled anti-mouse IgG antibodies. Between each step, theslides were washed with 1% BSA (Sigma) in PBS. Cells were examined usingconfocal fluorescence microscopy (Confocal 1024 microscope, Olympus AX70,Olympus Optical, Tokyo).

Teratoma formation and histological analysis

Eight-week-old nude mice (Samtako Bio Korea, Seoul, Korea) were anaesthetisedwith diethyl ether. Fifty microliters of a P15 DPPSC or DPMSC cell suspension(46107 cells/ml), from three different donors (14-, 17-, 28-year-old donors), mixedwith 50 ml of Matrigel (BD) was injected subcutaneously into the dorsal flanks ofthe mice, which were then housed with free access to water and food under specificpathogen-free conditions. After 3 or 5 weeks, the teratoma-like structures weresurgically dissected from the mice followed by fixation with 4% paraformaldehydeand 1.25% glutaraldehyde, and the mice were subjected to histological analysis.Specimens were embedded in paraffin, cut into 3 mm sections and stained withhaematoxylin and eosin (H&E).

Immunohistochemistry

Samples were cut into 7-mm-thick sections for immunohistochemical staining.Slides were placed in PBS for 30 minutes to remove gelatin. After being washedtwice with distilled water for 5 min, the sections were blocked against endogenousperoxidase in 0.3% hydrogen peroxide for 15 minutes and 10% normal goat serumin PBS for 1 hour at room temperature to reduce nonspecific antibody interactions.The slides were incubated with mouse monoclonal primary antibodies againsthuman OCT3/4 (1:400, BD), NANOG (1:400, BD), SSEA4 (1:400, BD) andLIN28 (1:400, BD) at 4 C overnight. After washing with PBS, the specimens wereincubated with biotinylated goat anti-mouse secondary antibody (Zymed) andstreptavidin peroxidase (Zymed) at room temperature for 10 minutes each. Finally,the specimens were visualised using a diaminobenzidine reagent kit (Zymed). Theimmunostained sections were counterstained with H&E.

Transmission electronic microscopy

A piece of the cell pellet measuring 1 mm3 was fixed in a solution of 2%formaldehyde, 2.5% glutaraldehyde and Karnoski buffer with cacodylate (0.2 mol/l, pH 7.4). After 48 hours, the samples were soaked in araldite. The ultra-finesections were stained for contrast with citrate and then observed using anelectronic microscope (Zeiss EM900).

Scanning electron microscopy

For SEM analysis, samples were fixed in 2.5% glutaraldehyde (Ted Pella) in 0.1 MNa-cacodylate buffer (EMS, Electron Microscopy Sciences, Hatfield, PA)(pH 7.2) for 1 hour on ice. After fixation, the samples were treated with 1%osmium tetroxide (OsO4) for 1 hour. The samples were dehydrated in serialsolutions of acetone (30–100%) with the scaffolds mounted on aluminium stubs.The samples were then examined using a Zeiss 940 DSM scanning electronmicroscope.

Cell migration assay

The cell migration capacity of DPPSCs and DPMSCs from the same donor, as wellas hNTERA cells, which were used as a positive control, was tested using aMultiscreen-MIC Plate polycarbonate filter (8 mM pore size) (Millipore).Approximately 16103 cells in 150 ml were added to the bottom wells of thefilter plate and incubated at 37 C in 5% CO2 for 6 hours. After incubation, thefilters were removed and the topside of the membrane was scraped to remove non-migrated cells. The filters were then stained with Toluidine Blue. The cells werecounted using a light microscope. Experiments were performed in triplicate andthe data were pooled.

Short-chromosome genomic hybridisation (short-CGH)

DPPSCs were isolated one by one by manual catching and short-CGH techniquewas developed (Rius et al., 2010) (n515). Hybridisation of control and DPPSCsamples was carried out against a masculine cell preparation.

Mesoderm differentiation

For bone differentiation, cells were seeded in six-well plates and in a Cell Carrier3D glass scaffold (Orla protein) on a 24-well plate with culture medium at adensity of 36103 cells P15 per cm2. After 24 hours, differentiation was initiated bythe following medium: a-MEM containing 10% heat inactivated FBS, 10 mM b-glycerol phosphate (Sigma), 50 mM of L-ascorbic acid (Sigma), 0.01 mMdexamethasone and 1% penicillin and streptomycin. The medium was changedevery 3 days for 21 days. Differentiated cultures were evaluated by qRT-PCR forALP, OSTEONECTIN, OSTEOCALCIN, OSTEOPONTIN, COLLAGEN I,COLLAGEN III, BMP-2 and NANOG every week. The cultures were alsoanalysed via immunofluorescence for OSTEOCALCIN and Alizarin Red stainingafter 3 weeks of differentiation.

Vessel-derived endothelial cell differentiation

Undifferentiated DPPSCs were cultured at a density of 36103 cells per cm2 withbasal media for 1 day. Culture medium was then exchanged for differentiationmedia (Basal Media, FBS 2%, VEGF 50 ng/mL, bFGF 10 ng/ml). Cells weregrown on coverslips and 6-well plates treated with Fibronectin and incubated at

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37 C at 5% CO2. The medium was changed every 3 days for 21 days.Differentiated cultures were evaluated by qRT-PCR and immunofluorescence.RNA was obtained at day 0 and then weekly and tested for endothelial gene Flk1and CD14 expression by qRT-PCR.

Endoderm differentiation

For 2D differentiation, DPPSCs P15 were seeded in 6-well plates with culturemedium at a density of 56104 cells/cm2. The following day, the culture mediumwas exchanged for differentiation medium consisting of base medium containing100 ng/ml HGF and 100 ng/ml FGF-4 (R&D Systems). The medium was changedevery 3 days for 3 weeks. Differentiated cultures were evaluated by qRT-PCRfor GATA4, HNF3b, HNF6, AFP, ALB, CEBPA and NANOG expression.Differentiated cells were identified using immunofluorescence microscopy foralbumin protein expression and albumin secretion analysis at different days ofdifferentiation.

For 3D differentiation, 56104 cells P15 were seeded in a Cell Carrier 3D glassscaffold (Orla protein) pre-coated with 2% Matrigel and placed in 24-well plateswith RPMI medium (Mediatech) supplemented with GlutaMAX and penicillin/streptomycin and containing 0.5% defined foetal bovine serum (FBS; HyClone)and 100 ng/ml Activin A (R&D Systems). Three days post-induction, the mediumwas refreshed using the same RPMI-based medium with 100 ng/ml Activin A butreplacing FBS by KOSR 2%. After 2 days, definitive endoderm cultures wererefreshed with RPMI medium supplemented with GlutaMAX and penicillin/streptomycin and containing 2% KOSR, 10 ng/ml FGF-4 (R&D Systems) and10 ng/ml HGF (R&D Systems). Three days later, the cells were switched tominimal MDBK-MM medium (Sigma-Aldrich) supplemented with GlutaMAXand penicillin/streptomycin and containing 0.5 mg/ml bovine serum albumin(BSA) (Sigma-Aldrich), 10 ng/ml FGF-4 and 10 ng/ml HGF. After another 3 days,the cells were switched to complete hepatocyte culture medium (HCM)supplemented with SingleQuots (Lonza) and containing 10 ng/ml FGF-4, 10 ng/ml HGF, 10 ng/ml oncostatin M (R&D Systems) and 1027 M dexamethasone(Sigma-Aldrich). Differentiation was continued for another 9 days. At each stage,the medium was refreshed every 2–3 days.

Neuroectoderm differentiation

For neural differentiation, cells P15 were seeded in 6-well plates and in 75 cm2

flasks in base culture medium at a density of 36103/cm2. The following day, theculture medium was exchanged for differentiation medium; the differentiationmedium differed from week to week. During the first week, the medium consistedof base medium and bFGF (100 ng/ml). During the second week, the mediumconsisted of base medium, FGF-8 (10 ng/ml) and SHH (100 ng/ml). During thethird week, the medium consisted of base medium, BDNF (10 ng/ml) and GDNF(10 ng/ml) + N2 (R&D Systems). The medium was changed every 3 days. Neuraldifferentiation was evaluated via qRT-PCR for TAU, NURR1, NESTIN, GFAP,MBP, SOX1 and NANOG expression. Cultures were also analysed byimmunofluorescence for PSD95-FITC and GALC-595 protein expression.

Alizarin Red staining

Cells were fixed in a 2.5% glutaraldehyde mixture that was freshly prepared in 16PBS buffer for 10–15 minutes at room temperature. Cells were then washed with16 PBS and 2% Alizarin Red solution (Millipore) was added to the fixed cells.Following incubation at 37 C for 20 minutes, cells were then observed throughmicrophotography (positively stained nodules are in orange-red).

Albumin secretion

The production of albumin was determined using the Albumin Assay Kit (Sigma)according to the manufacturer’s instructions.

Alkaline phosphatase staining

For alkaline phosphatase (ALP) staining, EBs were fixed in a solution of 4%paraformaldehyde in PBS for 20 minutes. After extensive washing in PBS, cellswere incubated in NTMT solution [10 mM NaCl, 100 mM Tris-HCl (pH 9),50 mM MgCl2, supplemented with 0.1% Tween-20] for 5 minutes and then inNTMT solution supplemented with NBT (Nitro-Blue Tetrazolium Chloride) andBCIP (5-Bromo-4-Chloro-39-Indolyphosphate p-Toluidine Salt) in darkness untilthe staining developed.

ALP activity

During the 3D osteoblast differentiation of DPPSCs and from passage number 15,ALP activity was quantified every week by spectrophotometry using a Cromatestkit (Linear), in accordance with the manufacturer’s instructions. We measured theabsorbance of each sample at 1, 2, 3, 5 and 10 minutes.

Calcium quantification

Differentiated cells were washed twice with 16 PBS. Accumulated calcium wasremoved from the cellular components using lysis solutions contained in the Sigma

kit for the analysis of calcium accumulation, according to the manufacturer’sinstructions. The total calcium was calculated using standard solutions and theabsorbance was measured at 575 nm.

Cytochrome P450 3A4 metabolic activity assay

Cytochrome P450 (CYP) 3A4 enzyme activity assay was assessed by measurementof luciferase activity with the P450-Glo CYP3A4 assay, according to themanufacturer’s instructions. Differentiated cells were incubated at 37 C in DMSOplus ethanol supplemented with 50 mmol/l luciferin PFBE (150 ml/well) andwithout DMSO; undifferentiated DPPSCs were used as negative control. After 3hours of incubation, 50 ml of medium was transferred in a 48-well plate and mixedwith 50 ml of luciferin detection reagent to initiate luminescent reaction. After20 minutes of incubation at room temperature, luminescence was measured with aVictor3 luminometer (PerkinElmer).

Statistical analysis

To assess the percentages of specific markers for DPPSCs, data were subjected to aregression analysis, which considered the independent variable (age) and thedependent variable (different markers). We established statistical significance at aP value less than 0.1 (90% confidence level). For all other data, the statistical testapplied was the paired samples t-test, with statistical significance set at P,0.05.Data were analysed with SPSS Version 16.0 software. The values are expressed asthe mean 6 s.d.

Ethical regulations

Dental pulp tissues used for these experiments were obtained with informedconsent from donors. All experiments were performed in accordance with theguidelines on human stem cell research issued by the Committee on Bioethics ofthe International University of Catalonia.

AcknowledgementsWe thank M. Costa for help with FACS analysis, as well as J.Navarro and J. del Rey for their dedication in cytogenetic analysisusing a newly developed CGH technique was performed in the Unitatde Biologia Cellular i Genetica Medica Eugin-UAB. In memory ofNuria Durany, without whom this article would not have beenpossible. Author contributions were as follows: A.M., conceptionand design, collection and assembly of data, data analysis andinterpretation, manuscript writing and final approval of themanuscript; C.G.-R., collection and assembly of data andmanuscript writing; M.F., collection and assembly of data; D.G.-F.,collection and assembly of data; M.B., data analysis andinterpretation; M.C., collection and assembly of data; H.-S.J,financial support, administrative support, data analysis andinterpretation; F.H.-A., provision of study patients; N.C., dataanalysis and interpretation; F.P., administrative support, and dataanalysis and interpretation; E.F.P., provision of study patients; L.G.,financial support, administrative support and final approval ofmanuscript.

FundingThe Universitat International de Catalunya supported all this work.This research received no specific grant from any funding agency inthe public, commercial or not-for-profit sectors.

Supplementary material available online at

http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.096537/-/DC1

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